Micrococcus luteus in a Veterinary Clinical Setting

Micrococcus luteus in a Veterinary Clinical Setting

Erin Murray

Biol 342, Section F02

 

Introduction

In veterinary clinics or hospitals, it is highly important to maintain as clean an environment as possible for the sake of their patients’ health. Failure to do so carries the possibility of inadvertently transferring bacterial or viral pathogens from one patient to the next, which could result in a healthy patient becoming sick simply from having visited the clinic. This is known as a nosocomial infection (Stull and Weese). If a patient is hospitalized and acquires a nosocomial infection, this could lead to its hospital stay being prolonged (Harper et al.). Microbes known to cause nosocomial infections have been isolated from samples taken from various surfaces around veterinary clinics, including medical equipment used on patients, as well as from air samples from the various rooms of an animal hospital (Harper et al.). This means that airborne bacteria could potentially land on and colonize surfaces of medical equipment even after they have been thoroughly cleaned.

In the study of airborne sampling, the most commonly isolated bacteria belonged to the genus Micrococcus (Harper et al.). Micrococcus species commonly grow on Mammalian skin as a commensal bacteria, and are rarely ever pathogenic. There have been rare cases where patients developed infections from Micrococcus bacteria becoming an opportunistic pathogen, but this has only occurred in patients that were immunocompromised (Hanafy et al., Public Health England).

The purpose of this study was to attempt to determine if any pathogenic bacteria were taking up residence on the surfaces of the medical equipment at a local emergency veterinary clinic. Speaking from personal experience, one potential source of contact transfer of microbes is a veterinarian’s or technician’s stethoscope. Though it is used on every patient walking through the doors of the clinic, the cleaning of this piece of medical equipment can be easily overlooked. Many veterinary staff will wear their stethoscope around their neck in order to keep it close at hand and may not get a chance to, or may even forget to, disinfect it in between patients.

I hypothesized that I would find species of bacteria that typically lived either on Mammalian skin and hair, or in canine or feline mouths. I suspected that I might find mouth bacteria because dogs and cats lick their fur to groom themselves. From this hypothesis I predicted that I would isolate a bacterium that could at least tolerate oxygen and would therefore be an aerobe or an aerotolerant anaerobe.

 

Methods

The diaphragm of a frequently used stethoscope at a local veterinary clinic was swabbed with a sterile cotton swab that had been wetted with sterile water. The swab was then used to inoculate a Tryptic Soy Agar (TSA) plate. The plate was sealed with parafilm and was stored at room temperature for one week to allow for bacterial colonies to grow. At the end of one week, several colonies of different colors and textures had grown, and one colony was selected to be isolated. The colony was goldenrod yellow in color, circular and raised, had a smooth and shiny finish, and was about the size of a pencil eraser. A sample of the colony was transferred onto a new TSA plate using the quadrant streak technique as outlined in the handout for Lab 2: Aseptic Technique. The plate was incubated at 37 °C to accelerate colony growth. A new quadrant streak was performed every 3-4 days for a total of 4 times to ensure that purity of the culture had been achieved.

A series of physiological tests were performed on the isolate to determine its physiological characteristics. A Gram stain was performed on a microscope slide of the culture, following the protocol as outlined in Lab 4: Staining Techniques, to determine the type of cell wall the bacteria possessed. Following the protocols as outlined in Lab 6: Physiological Testing of Your Isolate, a fluid thioglycollate test was performed to determine the culture’s oxygen class, an oxidase test was performed to determine whether the isolate produced the cytochrome c oxidase enzyme, and a catalase test was performed to determine whether the isolate produced the catalase enzyme. An API 20E test strip was performed, following the protocol as outlined in Lab 6, to determine which materials the bacteria could metabolize. The test strip was rechecked after 24-48 hours of incubation at 37 °C. Later on, an API Staph test strip was also performed, following the protocol on the API Staph handout. It, too, was incubated at 37 °C and rechecked 24-48 hours later.

The culture’s susceptibility and/or resistance to various antibiotics was tested, following the protocol as outlined in Lab 9. The antibiotics used for the test were amikacin, cefazolin, clindamycin, erythromycin, gentamicin, oxacillin, tetracycline, and tobramycin.

DNA was extracted from the isolate for genomic analysis, using the PowerSoil DNA isolation kit and following the protocol as outlined in the handout for Lab 5. The DNA sample was then submitted to the UAF DNA Core Lab to be sequenced. When the raw genomic data had been obtained, it was then analyzed on BaseSpace, using the SPAdes Genome Assembler, Kraken Metagenomics, and Prokka Genome Annotation programs. The contig data was then analyzed using the NCBI Nucleotide BLAST program.

 

Results

Gram staining of the bacterium revealed that the isolate was a Gram-positive cocci that formed tetrads and irregular clusters. In fluid thioglycollate the bacteria grew directly on and just below the surface of the fluid, and no growth was observed in the anoxic layer of the fluid, therefore the bacterium is likely a strict anaerobe. The isolate tested positive for the presence of the enzymes catalase and cytochrome c oxidase. The API 20E and API Staph assays had no positive results. Antibiotic resistance testing revealed that the isolate was susceptible to amikacin, cefazolin, clindamycin, gentamicin, tetracycline, and tobramycin, was intermediately susceptible to oxacillin, and was resistant to erythromycin.

SPAdes Genome Assembler discovered 300 contigs with the largest contig being 61,411 base pairs (bp). The total length of the genome was determined to be 2.51 Mbp, and the GC content of the genome was determined to be 72.72%. Prokka Genome Annotation discovered 53 tRNAs, 0 rRNAs, 1 CRISPR gene, and 2270 CDs. The Kraken Metagenomics program took 173,940 reads. Of those reads, 55,285 were classified, resulting in 31.78% of total reads being classified. 25.85% of the total reads were classified to the species level, and the bacterium was identified as Micrococcus luteus. Analyzing contigs via BLAST revealed that the sequence from node 21 was a 97% match to the genomes of the NCTC 2665 and trpE16 strains of M. luteus.

 

Discussion

After extensive testing of the isolate, it has been concluded that the identity of the isolate is Micrococcus luteus.

M. luteus is described in the literature as an aerobic, catalase- and oxidase-positive, Gram-positive non-motile cocci which form tetrads, with colonies being yellow in pigment, convex and smooth in texture with regular edges (Public Health England, Whitman et al.). The physical appearance of colonies of the isolate grown on agar plates, and physiological tests performed on the isolate are in agreement with these descriptions. Genome length and GC content of the isolate that were reported in SPAdes also agree with values in the literature (Hanafy et al., Whitman et al.)

Neither of the API assays that were attempted had any color changes, meaning that none of the tests came back positive. For the API 20E test, the lack of positive results is likely due to the test being the wrong fit for this microbe – it was meant to be used on Gram-negative anaerobes, and the isolate was a Gram-positive strict aerobe. It is possible that the reagents were unable to penetrate the thick peptidoglycan layer of the bacteria’s cell walls in order for metabolism of the reagents to occur.

The API Staph test should have been the best fit for the isolate. However, it still came back with no positive results. Researching the literature reveals that M. luteus should be able to metabolize glucose, fructose, sucrose, and mannose (Hanafy et al.), meaning that the GLU, FRU, SAC, and MNE wells on the API Staph test should have had color changes to indicate positive results. Some strains have been found to also metabolize maltose and trehalose, contrary to prior data indicating that these molecules were not utilized by M. luteus (Wieser et al.), meaning that there possibly could have been positive results from the MAL and TRE wells if the isolate was a similar strain. The literature also indicates that M. luteus is urease positive (Whitman et al.), so the URE well should have indicated a positive result as well. The lack of results from the API Staph test was probably due to not having a fresh enough sample: the instructions for the test call for a sample that is 18-24 hours old, and the plate that the sample was taken from had been incubating for between 4 and 7 days.

The isolate was found to be susceptible to most of the antibiotics that were tested, except for erythromycin, which it was completely resistant to. Resistance to erythromycin has been found to be due to a plasmid present in a strain of M. luteus isolated from human skin (Liebl et al.). With humans and companion animals having such an intimate relationship and being in contact with each other so often, it doesn’t take a stretch of the imagination to suggest that erythromycin-resistant M. luteus could be transferred from human skin onto the skin of a companion animal, or vice versa, which would explain the presence of erythromycin-resistant M. luteus on a stethoscope in a veterinary clinic. Horizontal gene transfer of this resistance plasmid could potentially cause other bacterial species to develop resistance as well.

Further investigation into the types of bacteria growing on medical equipment in veterinary clinics should be pursued. Only one of several different colonies from the original swab of the stethoscope was chosen for further evaluation. Analysis of the other colonies that were present on the original TSA plate would be the first step in continuing the investigation, followed by taking samples from other objects and equipment, and possibly obtaining airborne samples from around the clinic. This type of investigation would be very helpful in assessing local clinics’ risk of exposing their patients to nosocomial infections.

 

Figures

Figure 1. Macroscopic and microscopic morphology of the isolate. Colonies are yellow, convex, and smooth in texture with a regular edge. Gram staining reveals that the bacteria are Gram-positive cocci that form tetrads and irregular clusters.

Figure 2. Antibiotic resistance testing of the isolate. The isolate showed resistance to erythromycin, but to no other antibiotic which was tested.

Figure 3.  Krona Classification chart from the Kraken Metagenomics program. The reads were classified and nested into successive taxonomic groups, down to the species level.

 

References

Hanafy R.A., Couger M.B., Baker K., Murphy C., O’Kane S.D., Budd C., French D.P., Hoff W.D., and Youssef N. 2016. Draft genome sequence of Micrococcus luteus strain O’Kane implicates metabolic versatility and the potential to degrade polyhydroxybutyrates. Genomics Data 9:148—153.

Harper T.A., Bridgewater S., Brown L., Pow-Brown P., Stewart-Johnson A., Adesiyun A.A. 2013. Bioaerosol sampling for airborne bacteria in a small animal veterinary teaching hospital. Infection Ecology and Epidemiology 3.1.

Liebl W., Kloos W.E., and Ludwig W. 2002. Plasmid-borne macrolide resistance in Micrococcus luteus. Microbiology 148:2479—2487.

Public Health England 2014. Identification of Staphylococcus species, Micrococcus species and Rothia species. Bacteriology — Identification 3:1-32.

Stull J.W., and Weese J.S. 2015. Hospital-Associated Infections in Small Animal Practice. Veterinary Clinics: Small Animal Practice 45:217—233.

Whitman M., Goodfellow M., Kämpfer P., Busse H.J., Trujillo M., Ludwig W., Suzuki K.I. 2012. Bergey’s Manual of Systematic Bacteriology: Volume 5: The Actinobacteria. Springer Science & Business Media, p. 574-575.

Wieser M., Denner E.B.M., Kämpfer P., Schumann P., Tindall B., Steiner U., Vybiral D., Lubitz W., Maszenan A.M., Patel B.K.C., Seviour R.J., Radax C., and Busse H.J. 2002.  Emended descriptions of the genus Micrococcus, Micrococcus luteus (Cohn 1872) and Micrococcus lylae (Kloos et al. 1974). International Journal of Systematic and Evolutionary Microbiology 52:629—637.

Cheek Swab Reveals Presence of Stapholococcus aureus in Oral Cavity of some Individuals

Cheek Swab Reveals Presence of Stapholococcus aureus in Oral Cavity of some Individuals

Mike Fierro

Uaf undergraduate and BIOL 342 student

 

Introduction:

From childhood we learn about the different environments around the world ranging from the tundra of the arctic to the arid sands of the desserts; with further research we discover the animals that are able to thrive in these environments through adaptations to the conditions.   It’s easy to focus on a narrow range of environments to ones we are able to encounter through exploration, and to limit our knowledge of organisms to animals we may come across in those areas we can reach.   But what about environments that we don’t think about as being environments? Environments aren’t always large tracts of land or a lake system.   What if the area a creature inhabits is unlike any you will ever come across and yet at the same time you are more connected to than any place that you could find on a travel?   What if the environment… is you?

With over 700 bacterial species known, the human oral cavity is a perfect home for many bacteria.   Bacteria are able to grow on several sites in the mouth including the tongue dorsum, lateral sides of tongue, buccal epithelium, hard palate, soft palate, supragingival plaque of tooth surfaces, subgingival plaque, maxillary anterior vestibule, and tonsils (Aas et al. 2005).   There are numerous benign as well as opportunistic pathogens that inhabit the oral cavity as well including Staphylococcus aureus, Streptococcus mitis, and Steptococcus pyogenes as well as species of the yeast Candida (Lab 10).   The objective of this study to isolate a bacteria and perform both physiological and genetic tests to determine its identity as well as how and where it survives.   With a simple swabbing of the inside my cheek, I am making the attempt to find an organism that calls the human oral cavity home.

Methods:

A sterile swab was used to rub several times against the inside of my cheek, avoiding other areas of the mouth.   The sample was quickly inoculated onto two microbial growth media mediums: a tryptic soy agar (TSA) plate, as well as a Sabouraud’s agar (SA) plate, where it was allowed to grow at around 23 ºC for several days.

In lab the original TSA sample was kept and four different quadrant streaks were carried out throughout the next couple weeks in order to purify the isolate.   After these attempts to purify the culture, a series of physiological and genetic tests were used in order to obtain an identity of the bacteria isolated from the cheek swab.   The physical tests included a Gram stain, fluid thioglycollate test, oxidase test, catalase test, and an API-20E test, while DNA purification followed by a BaseSpace analysis was used as a genetic test of the isolate.

Three weeks into the project, the most recent TSA plate was used in order to perform a Gram stain (Lab 4 protocol), which would identify if the cell wall of the bacteria had a thick (positive Gram stain) or thin (negative Gram stain) peptidoglycan layer.   A positive Gram stain dyes the cells a dark purple due to crystal violet being encased in the cell wall due to dehydration of the peptidoglycan layer followed by addition of Safranin, whereas crystal violet is removed by ethanol from a Gram negative cell and the cell turns a light pinkish red after the introduction of Safranin.

Five weeks from original sampling, three physiological tests were performed on a fresh culture sample.   The fluid thioglycollate test was carried out (Lab 6 protocol) in order to determine the oxygen class of the isolate.   The solidified agar is a barrier to oxygen, so the bottom of the test tube is anoxic while the top has oxygen available.   This test reveals if the bacteria is able to grow only in the presence of oxygen (strict aerobe), grow regardless but better with oxygen (facultative aerobe), grow just below the surface (microaerophile), only grow without oxygen’s presence (strict anaerobe), or equally throughout (aerotolerant anaerobe).   The oxidase test was used (Lab 6 protocol) to determine the presence or lack of cytochrome oxidase, which differentiates bacteria into pseudomonad species (oxidase positive) and enteric species (oxidase negative).   The catalase test reveals the presence of the catalase enzyme, which protects cells that have it from reactive oxygen species including hydrogen peroxide.   By neutralizing these oxygen species, the cell is protected from oxidative damage.   The API-20E test is comprised of twenty physiological tests which characterize the Gram negative bacteria.   The characteristics tested are an indication of what the bacteria is able to ferment as well as production of certain enzymes and ability to oxidize reactants.

A DNA extraction was performed (Lab 5 protocol) in order to obtain the genomic sequence of the bacterial isolate.   The extraction is carried out in three main steps: cell lysis, removal of inhibitors and proteins, and obtaining a pure solution of DNA.   Cell lysis is accomplished by physical means of using beads to batter the sample, chemical means to dissolve the cell wall, and by enzymes that break down chunks of cell wall, which allow the cell’s contents to be exposed.   Proteins and inhibitors are used by introducing chemical substances that extract these from the solution followed by centrifugation.   During the centrifugation, the DNA is bound to a solid matrix while the inhibitors and proteins are part of the supernatant.   In the last step, DNA is removed from the solid matrix, centrifuged again, and appears in the resulting supernatant.

The last test on the isolate performed was to determine its antibiotic susceptibility (Lab 9 protocol).   Seven antibiotic discs in total were placed among the isolated culture to determine if zones of inhibition were produced by the antibiotics that prevented its bacterial growth.   The antibiotics tested included tobramycin, cefazolin, vancomycin, cefoperazone, trimethoprim, gentamicin, and amikacin.   The results were compared to known susceptibility thresholds of each antibiotics in order to determine efficacy of the antibiotic.

Results:

After the first 24-hour period after inoculation, no colonies were evident on either plate.   After 72 hours, numerous colonies of varying sizes were forming on the TSA plate compared to one large colony on the SA plate.   After five days, the number of colonies on the TSA plate had reached double digits while the SA plate still had one major colony with a couple smaller colonies on the side.   After one week’s time, the TSA plate had around 30 colonies; there was one large colony, four colonies larger than 1mm, and the amount of pencil-tip sized colonies was in the double digits.   On the SA plate, however, there was no noticeable growth than observed on day 5.

Figure 1. A comparison between colonies formed on SA plate (left) and TSA plate (right).

The results of the physiological tests will be discussed first.   Under the microscope, there was Gram negative bacteria along with Gram positive bacteria.   The sample was continuously purified thereafter until the resulting bacteria was completely Gram positive.   The fluid thioglycollate test revealed that growth occurred more on the surface than the underlying agar.   The oxidative test on the bacterial isolate was a yellow/buff color, which occurs when the test is negative.   The catalase test showed that the isolated bacterial bubbled when exposed to hydrogen peroxide.   This is a positive result to the catalase test.   The original API-20E test revealed that the bacterial contaminant did not have gelatinase and thus couldn’t break down gel, could oxidize nitrate to nitrite, had a positive citrate test, and was able to ferment glucose, mannitol, inositol, sorbitol, rhamnose, sucrose, meliblose, amygdalin, and arabinose.   The bacteria had large zones of inhibition on all antibiotics tested.

Figure 2. Picture of contaminated Gram stain.                         Figure 3. Example of zone of inhibition.

Figure 4. Picture of thioglycollate test.                                       Figure 5. Picture of contaminant API-20E test.

 

 

The genetic results showed that the number of contigs in the sample tested was in the hundreds, the total length was in the millions, and the largest contig was over 100,000 bp.   The result of the BaseSpace analysis was that 97% of all contigs were able to be read; of those contigs, there was a 99.94% match with known samples of Staphylococcus aureus.

Figure 6.   Pie diagram and bar graph results of BaseSpace test revealing Staphylococcus aureus as isolate.

Discussion:

                      TSA favors bacterial growth while SA provides a better medium for fungal growth to thrive.   The reason more colonies grew on the TSA plate was because more colonies were bacterial and made more use of the medium that favored them.   The reason colonies existed on the SA plate was because either there was a fungus within the cheek, or there was bacteria that was able to survive with the resources available in the SA plate.

The original Gram stain result of both Gram positive and Gram negative bacteria proves the bacterial sample was not pure.   A purified sample of bacteria would be one or the other, unless the bacteria is Gram variable, which is very rare and did not match the known scientific literature with the results of the genomic sequencing.   The gram positive result on the culture after further purification did agree with scientific literature.   Under the microscope, the bactieria formed clusters of round cells.

The fluid thioglycollate test shows the bacteria is a facultative anaerobe as it grows more when oxygen is available, but is able to survive when oxygen is not present.     The negative result on the oxidase test reveals the bacterial isolate is an enteric species, while the positive catalase test reveals that the culture has a resistance to reactive oxygen species and thus protects the cell from oxidative damage The API-20E test results showed that there was a Gram negative bacteria present in the culture that was not present in later Gram stains.   Unfortunately, an API-Staph was not carried out since the writer of this paper missed the part about API-20E working on Gram negative bacteria only when he had decided the isolate was Gram positive.   The antibiotic disc test revealed the isolate is highly susceptible to all seven antibiotics tested when compared to their susceptibility thresholds.

The number of contigs, total length, and the size of the largest contig were all statistically meaningful (Lab 7 protocol).   The high amount of readable contigs as well as the nearly identical comparisons between laboratory results and scientific literature suggest to the utmost that the bacteria isolated from the cheek swab is Staphylococcus aureus.

Using the API-20E test, the Gram negative result, and the observations of the bacterial contaminant under a microscope, it would be possible for future research to determine the identity of this contaminant.   It is interesting that the contaminant yielded a catalase positive, oxidative negative, and citrus positive result, all of which are shared by the Gram positive Staphylococcus aureus that was isolated.   I attribute this to the similarity of the environment from which these bacteria were isolated.

The remainder of this paper will discuss the bacterial isolate.   Staphylococcus aureus is found in clusters of round cells in the nasal passages, oral cavity, lower reproductive tract, and skin of humans as well as other animals such as the dogs of healthcare workers (Masalha et al. 2001).   It’s estimated that 20-30% of people are long term carriers (Tong et al. 2015).     Usually a commensal bacteria, it becomes an opportunistic pathogen when circumstances allow and cause ailments ranging from minor skin infects and food poisoning to life-threatening diseases such as pneumonia, endocarditis, toxic shock, and sepsis (MedlinePlus 2017).   Once it’s inside the bloodstream, Staphylococcus aureus has enzymes to clot blood and destroy proteins, can secrete toxins, and has super-antigens that cause organ failure by causing the immune system to become hyper-reactive.   Antibiotic-resistant strains such as methicillin-resistant Staphylococcus aureus (MSRA) are becoming increasingly abundant and are a major concern in the medical field due to transmissions in hospitals.   Over half a million people in US hospitals contract Staph infections, and most are caused by Staphylococcus aureus (Bowersox 1999).   Staphylococcus aureus is here to stay; it would benefit us to further research this bacteria to better understand it to aid in limiting its role as an opportunistic pathogen that has taken a countless amount of lives.

 

 

Works Cited:

Aas J; et al. (2005). “Defining the Normal Caterial Flora of the Oral Cavity’. J Clin Microbiol. November 2005. 43(11) 5721-5732.

Bowersox, John (27 May 1999).  “Experimental Staph Vaccine Broadly Protective in Animal Studies”. NIH. Archived from  the original  on 5 May 2007. Retrieved  28 July  2007.

Masalha M; et al. (2001).  “Analysis of Transcription of the Staphylococcus Aureus Aerobic Class Ib and Anaerobic Class III Ribonucleotide Reductase Genes in Response to Oxygen”.  Journal of Bacteriology.  183  (24): 7260—7272.

MedlinePlus [Internet]. “Staphylococcal Infections”. Bethesda, MD: National Library of Medicine, USA.  Skin infections are the most common. They can look like pimples or boils.

Tong SY; Davis JS; Eichenberger E; Holland TL; Fowler VG (July 2015).  “Staphylococcus aureus infections: epidemiology, pathophysiology, clinical manifestations, and management”

UAF Lab handouts provided to me by Mary Beth Leigh.

The Microbiology of Music

 

The Microbiology of Music – Describing a Microbe Found in a Flute Headjoint

Chaya Pike

Biol 342, Spring 2017

 

Introduction

It has long been the practice of well-meaning parents to encourage their offspring to pick up a musical instrument, in the hopes that it will make them smarter, more disciplined, or more attractive to selective schools. Whether or not any of these hopes are realized, few parents likely recognize the potential dangers lurking within the wind instruments their children play. Several authors have documented respiratory infections, some of them serious, contracted from saxophones (Lodha & Sharma, 1988; Metzger et al., 2010), trombones (Metersky et al., 2010), and other woodwind and brass instruments (Glass, Conrad, Kohler, & Bullard, 2011; Marshall & Levy, 2011; Rackley & Meltzer, 2011). While not all microorganisms are inherently pathogenic to humans, the warm, moist (and rarely cleaned) interior surfaces of wind instruments provide a prime environment for a wide variety of pathogenic and non-pathogenic microbes to grow.

The objective of this paper is to isolate, characterize, and identify a slow-growing bacterium collected from the interior surface of a metal flute headjoint. Methods used to identify the bacterium include a battery of physiological testing and genomic sequencing.

Intriguingly, this microbe was one of only two from the sample capable of growing under room-temperature conditions on TSA. This is consistent with previous studies that have found reduced microbial loads on non-reed wind instruments compared to reed instruments (Marshall & Levy, 2011).

Methods

Sample collection and isolation

Two damp, sterile swabs were used to collect microbial samples from the inside surface of a metal flute headjoint, just below the embouchure hole. The flute had been played but not cleaned approximately twenty-four hours previously. The swabs were then used to inoculate a trypticase soy agar (TSA) plate which was then incubated at room temperature for five days.

Once visible colonies had formed, a single colony from the TSA plate was used to inoculate a fresh TSA plate using the streak plate method. This was repeated once more to obtain a pure culture.

Morphological and physiological characterization

The pure culture was examined for details about colony size, color, shape, distribution, and other relevant colony morphological characteristics. A Gram stain was performed on the isolate using the procedure outlined in Lab Handout 4, and the stained isolate was examined under the microscope for the aforementioned morphological characteristics, as well as to determine if the isolate was Gram positive or Gram negative.

Catalase and oxidase tests were performed to determine the enzymes used by the isolate in the electron transport chain, and a suite of 21 physiological tests (including tests for fermentation of glucose, mannitol, sorbitol, and others) were performed using an API 20E test strip. Protocols for each of these tests, described in Lab Handout 6, was followed exactly.

DNA extraction and analysis

A trypticase soy broth was inoculated with the isolate, but did not yield enough cells to perform a proper genomic DNA extraction, so additional colonies from an agar plate were added to the broth culture immediately prior to DNA extraction. Excepting this detail, genomic DNA extraction protocol described in Lab Handout 5 was followed. Using a PowerSoil DNA kit, DNA was isolated from the liquid culture and sequenced using the Illumina MiSeq DNA sequencer.

Genomic data produced during DNA sequencing was analyzed using Illumina BaseSpace. The genome was assembled using the SPAdes Genomic Assembler tool, taxonomic assignments were made using the Kraken Metagenomics tool, and functional genes were identified using the Prokka Gene Annotation tool. Additional analyses were performed using BLAST nucleotide alignment, and Dr. Eric Collins conducted analysis using Bandage software to assemble the isolate genome.

Results

Morphology and physiological tests

The isolate colonies appeared to be perfectly circular, and quite small (<2mm diameter). They were a consistent matte coral-pink color, and projected slightly from the surface of the agar plate. The colonies were incredibly slow-growing, and it took three days for the initial environmental sample to demonstrate visible microbial growth. Growth rates did not appear to be affected by temperature, and the isolate was similarly slow-growing in 37oC incubator and 4oC  refrigerator alike. When viewed under the microscope, the cell morphology was difficult to discern – the cells could either be irregularly shaped and arranged in chains, or elongate with constrictions along each cell (Figure 1).

Figure 1: Gram-stained isolate under 1000x magnification

Results of the Gram-stain indicate that the isolate is Gram-negative. Tests for catalase and oxidase were also negative. Results from the API 20E test strip are shown in Table 1. The only positive physiological tests for this isolate were for arginine dihydrolase, gelatinase, and glucose metabolism. Arginine dihydrolase is present in cells that use arginine as a source of carbon and energy, gelatinase is used to break down gelatin into useful sub-compounds, and glucose fermentation enables cells to use glucose as a carbon source and produces acidic byproducts.

Table 1: Results of API 20 E test strip.

Subtest Reaction/Enzyme Result
ONPG beta-galactosidase negative
ADH arginine dihydrolase positive
LDC lysine decarboxylase negative
ODC ornithine decarboxylase negative
CIT citrate utilization negative
H2S H2S production negative
URE urease negative
TDA tryptophane deaminase negative
IND indole production negative
VP acetoin production negative
GEL gelatinase positive
GLU glucose fermentation/oxidation positive
MAN mannitol fermentation/oxidation negative
INO inositol fermentation/oxidation negative
SOR sorbitol fermentation/oxidation negative
RHA rhamnose fermentation/oxidation negative
SAC saccharose fermentation/oxidation negative
MEL melibiose fermentation/oxidation negative
AMY amygdalin fermentation/oxidation negative
ARA arabinose fermentation/oxidation negative

 

DNA analysis

Genome assembly software yielded 347 contigs of greater than 1000 bp in length from the isolate, with a maximum length of 45702 bp. Functional gene annotation software found 55 tRNAs within the isolate sample, and a total of 2197 coding genes. Metagenomic analysis classified the isolate as Micrococcus luteus, with 98.75% of reads classified to the species level (79.85% of analyzed reads).

As M. luteus is morphologically and physiologically unlike the isolate in question, contamination was suspected. Further bioinformatic analysis conducted by Dr. Eric Collins suggests the isolate is another species within the genus Micrococcus, and likely possesses the mercury(II) reductase gene. This gene enables the microbe to use elemental mercury, NADP+, and H+ as substrates for the generation of NADPH, with Hg2+ as a byproduct.

Discussion

The morphological, physiological, and genetic results of this study are inconsistent and inconclusive. Morphologically, the isolate cells bear some resemblance to fungal hyphae, but considering that nothing grew on the Sabouraud agar plate, which selects for fungi, this result is highly unlikely. Genetic sequencing suggests the isolate may be the species Micrococcus luteus, but morphological observations and physiological tests do not align with this species identification. M. luteus forms yellow or greenish-yellow colonies on plated cultures, and the cells are cocci arranged in tetrads, while the colonies of this isolate are pinkish-coral colored with irregularly shaped, elongate cells (Kocur, Pacova, & Martinec, 1972). Furthermore, M. luteus is a Gram-positive, oxidase-positive, gelatinase-positive species, while this isolate is Gram-negative, oxidase-negative, and gelatinase-positive (Kocur et al., 1972).

Additional bioinformatic analysis done by Alexis Walker suggest another possible species within the genus Micrococcus M. roseus is similar in color to the isolate, and is also oxidase-negative (Mohana, Thippeswamy, & Abhishek, 2013). However, both M. roseus and M. luteus are catalase positive and negative for arginine dihydrolase, which is inconsistent with the physiological test results of this isolate. A summary of morphological and biochemical characteristics for M. luteus, M. roseus, and this isolate appears in Table 2.

 

Characteristic M. luteus M. roseus Isolate
pigmentation yellow red pink/coral
cell morphology tetra head coccus tetra head coccus pleiomorphic/ undefined
Gram stain positive positive negative
catalase test positive positive negative
oxidase test positive negative negative
glucose fermentation negative positive positive
gelatin hydrolysis positive negative positive
arginine dihydrolase negative negative positive

Table 2. Morphological and biochemical characteristics of M. luteus, M. roseus, and experimental isolate (Government of Canada, 2011; Kocur et al., 1972; Mohana et al., 2013).

 

Bioinformatic analysis done by Dr. Eric Collins suggests that the genus Micrococcus may be a viable identity for this isolate, even if M. luteus and M. roseus are not. Species within the genus Micrococcus are ubiquitous, and can be found on human skin and any surfaces human skin has been in contact with, which potentially includes the surfaces of a musical instrument (Government of Canada, 2011). However, all known species of Micrococcus are Gram-positive, which this isolate is not (Government of Canada, 2011; Mohana et al., 2013).

Additionally, Dr. Collins’ analysis identified the mercury(II) reductase (merA) gene as a likely component of the isolate genome. The merA gene is part of the mer operon, which reduces Hg(II) to Hg(0) and operates optimally at a slightly basic pH and temperatures between 37℃ and 45℃ (Giovanella, Cabral, Bento, Gianello, & Camargo, 2016). This isolate was sampled from the inside of a flute headjoint composed of silver and nickel, both of which are known to have some inhibitory effect on bacterial growth (Argueta-Figueroa, Morales-Luckie, Scougall-Vilchis, & Olea-Mejía, 2014/8; Clement & Jarrett, 1994). It is possible that the isolate used a similar resistance mechanism to the merA gene to live in a silver and nickel enriched environment, but that mechanism was not found and is purely conjectural.

This isolate is extremely slow-growing, and ultimately did not survive in plated culture for more than seven weeks. As such, it was not possible to streak a sufficient number of plates to guarantee that the culture was pure, though it visually appeared free from contaminants. It is likely that M. luteus was a contaminant in the culture or was introduced as a contaminant during the DNA extraction process. Additionally, because the cultures used for physiological tests were two weeks old or older, the results of those tests may be invalid.

In conclusion, the results of morphological observations, physiological tests, and genomic sequencing are inconsistent, and these inconsistencies could be due to incorrect lab techniques, contamination, or other factors. In future, when characterizing this microbe, it would be useful to test different growth media and incubation temperatures, as TSA and 37℃ were clearly not favorable growing conditions for this organism. Additionally, it would be fruitful to extract and sequence several samples of DNA from multiple colonies, to increase the probability of identifying non-contaminant sequences.

Citations

Argueta-Figueroa, L., Morales-Luckie, R. A., Scougall-Vilchis, R. J., & Olea-Mejía, O. F. (2014/8). Synthesis, characterization and antibacterial activity of copper, nickel and bimetallic Cu—Ni nanoparticles for potential use in dental materials. Progress in Natural Science: Materials International, 24(4), 321—328.

Clement, J. L., & Jarrett, P. S. (1994). Antibacterial silver. Metal-Based Drugs, 1(5-6), 467—482.

Giovanella, P., Cabral, L., Bento, F. M., Gianello, C., & Camargo, F. A. O. (2016). Mercury (II) removal by resistant bacterial isolates and mercuric (II) reductase activity in a new strain of Pseudomonas sp. B50A. New Biotechnology, 33(1), 216—223.

Glass, R. T., Conrad, R. S., Kohler, G. A., & Bullard, J. W. (2011). Evaluation of the microbial flora found in woodwind and brass instruments and their potential to transmit diseases. General Dentistry, 59(2), 100—7; quiz 108—9.

Government of Canada, P. H. A. of C. (2011, April 19). Micrococcus Pathogen Safety Data Sheet. Retrieved April 12, 2017, from https://www.phac-aspc.gc.ca/lab-bio/res/psds-ftss/micrococcus-eng.php#endnote8

Kocur, M., Pacova, Z., & Martinec, T. (1972). Taxonomic Status of Micrococcus luteus (Schroeter 1872) Cohn 1872, and Designation of the Neotype Strain. International Journal of Systematic Bacteriology, 22(4), 218—223.

Lodha, S., & Sharma, O. P. (1988). Hypersensitivity pneumonitis in a saxophone player. Chest, 93(6), 1322.

Marshall, B., & Levy, S. (2011). Microbial contamination of musical wind instruments. International Journal of Environmental Health Research, 21(4), 275—285.

Metersky, M. L., Bean, S. B., Meyer, J. D., Mutambudzi, M., Brown-Elliott, B. A., Wechsler, M. E., & Wallace, R. J., Jr. (2010). Trombone player’s lung: a probable new cause of hypersensitivity pneumonitis. Chest, 138(3), 754—756.

Metzger, F., Haccuria, A., Reboux, G., Nolard, N., Dalphin, J.-C., & De Vuyst, P. (2010). Hypersensitivity pneumonitis due to molds in a saxophone player. Chest, 138(3), 724—726.

Mohana, D. C., Thippeswamy, S., & Abhishek, R. U. (2013). Antioxidant, antibacterial, and ultraviolet-protective properties of carotenoids isolated from Micrococcus spp. Radiation Protection and Environment, 36(4), 168.

Rackley, C. R., & Meltzer, E. B. (2011). Throw caution to the wind instruments. Chest, 139(3), 729; author reply 729—30.